Plastids, the major organelles found only in plant and algal cells, are responsible for photosynthesis, for the storage of a wide variety of products, and for the synthesis of key molecules required for basic structural and functional aspects of plant cells. For example, plastids are responsible for the biosynthesis of purines and pyrimidines, and are the sole site of the synthesis of chlorophylls, carotenoids, certain amino acids (the “essential” amino acids), starches, fatty acids, and certain lipids.
Plastids are derived from proplastids, which are always present in young meristematic regions of a plant (a meristem is an undifferentiated region from which new cells arise). Proplastids can give rise to several different types of plastids, which types include: amyloplasts, unpigmented plastids which contain starch granules and which are especially common in storage organs, such as potato tubers; leucoplasts, colorless plastids involved in the synthesis of monoterpenes, the volatile compounds contained in essential oils and many of which are of commercial importance; chloroplasts, the green photosynthetic plastids responsible for energy capture via photosynthesis; and chromoplasts, yellow, orange, or red plastids, depending upon the particular combination of carotenes and xanthopylls present, and which are responsible for the colors of many fruits (tomatoes, oranges), flowers (buttercups, marigolds) and roots (carrots, sweet potatoes).
Plastids arise from the binary fission of existing plastids, independently of cell division. In root tips, shoots, and other meristems, proplastid division keeps pace with cell division, so the daughter cells possess approximately the same number of plastids as the parent cells; in angiosperms, this number is about 20 proplastids per cell. As cell expansion supersedes cell division, the number of plastids per cell increases due to continued plastid division. The number of plastids present in a mature plant cell is typically similar for a particular cell in a particular tissue; for example, an Arabidopsis leaf mesophyll cell typically contains about 120 chloroplasts. Thus, plastid division is essential for the maintenance of plastid populations in plant cells undergoing division, and for the accumulation of large chloroplast numbers in photosynthetic tissues.
Plastids are surrounded by a double membrane system which is made up of the outer and inner envelopes. The soluble interior portion of the plastid inside the inner envelope is the stroma; additional membrane structures may be present within the stroma, such as thylakoids. Thylakoids appear as interconnected stacked grana present in green chloroplasts, and contain the pigments necessary for light capture, such as chlorophyll. Thus, plastid division involves division of the outer and inner envelopes, as well as of the stroma and interior structures. As determined by ultra structural studies, plastid division begins with a constriction in the center of the plastid. Formation of the constriction is frequently associated with the appearance of an electron-dense annular structure termed the plastid dividing (PD) ring. In some electron micrographs of plastids from plants, the PD ring can be resolved into two concentric rings, an inner PD ring associated with the stromal surface of the inner envelope membrane, and an outer PD ring associated with the cytosolic surface of the outer envelope membrane. In other electron micrographs of plastids from red algae, yet a third PD ring is observed in the intermembrane space between the inner and outer envelope membranes. The constriction deepens and tightens, creating an extremely narrow isthmus before the two daughter plastids separate completely.
The mechanisms mediating plastid division are poorly understood, although it is believed that the PD rings are a dynamic macromolecular complex. It is also believed that this macromolecular complex is composed of numerous proteins that coordinate the mechanical activity required to constrict the plastid. Only a few components of the plastid division complex have been identified to date.
Plastid division is believed to have its evolutionary origin in a cyanobacterial endosymbiont that gave rise to chloroplasts (Osteryoung, K W et al. (1998) Plant Cell 10: 1991-2004). Thus, it has been proposed that the plastid division apparatus might have components in common with those involved in prokaryotic cell division, and in particular with cyanobacterial cell division (Possingham, J V and Lawrence M E (1983) Int. Rev. Cytol. 84: 1-56; and Suzuki, K et al (1994) J Cell Biol 63: 280-288). Genes from non-photosynthetic bacteria which play a role in division have been sequenced and identified. However, only a few of these genes involved in cyanobacterial division have been identified to date. One identified gene encodes bacterial FtsZ (from filamentation temperature-sensitive mutants, or fts mutants), which is a structural homologue to, and very likely the evolutionary precursor of, the eukaryotic tubulins (Erickson, H P (1998) Trends Cell Biol 7: 362-367; Faguy, D M and Doolittle W R (1998) Curr Biol 8: R338-341; Lowe, J and Amos L A (1998) Nature 391: 203-206) and Nogales, E et al. (1998) Nat Struct Biol 5: 451-458). FtsZ is well known to be a self-polymerizing, filament-forming GTPase, and it functions during bacterial cell division by assembling into a ring structure at the division site on the interior surface of the cytoplasmic membrane (Bi, E and Lutkenhaus J (1991) Nature 354: 161-164). The FtsZ ring assembly is required for the subsequent midcell localization of all other components of the cell division apparatus (Addinall, S G et al (1996) J Bacteriol 178: 3877-3884; and deBoer, P A J et al. (1988) J Bacteriol 170: 2106-2112); it remains associated with the leading edge of the division septum throughout cytokinesis, then it disassembles immediately following cell separation before rapidly reassembling at the center of the newly formed daughter cells (Addinall, S G et al (1996) J Bacteriol 178: 3877-3884; Bi, E and Lutkenhaus J (1991) Nature 354: 161-164; Butterfass, T (1988) in Division and Segregation of Organelles (Cambridge, UK; Cambridge University Press) pp 21-38; and Sun, Q and Margolin, W (1998) J Bacteriol 180: 2020-2056). In E. coli, placement of the FtsZ ring is governed by the minB operon, which encodes three gene products: MinC, MinD, and MinE (Lutkenhaus, J (1998) Curr Opin Microbiol 1: 210-215; Rothfield, L (1999) Annu Fev Genet 33: 423-448; Rothfield, L I and Justice, S S (1997) Cell 88: 581-584; and Sullivan, S M and Maddock, J R (2000) Curr Biol 10: R249-252).
FtsZ genes have also been found in nuclear genomes of land plants, as determined from plant gene database analysis. The encoded proteins fall into two major groups, FtsZ1 and FtsZ2 (Osteryoung K W, Stokes K D, Rutherford S M, Percival A L, and Lee, W Y (1998), Plant Cell 10: 1991-2004). FtsZ1 family proteins appear to contain cleavable chloroplast transit peptides at their amino terminal ends that target them to the chloroplast stromal compartment (Emanuelsson O, Nielsen H, Brunak S, von Heijne G (2000) J. Mol. Biol. 300:1005-16), whereas members of the FtsZ2 family do not appear to possess easily recognizable chloroplast transit sequences. However, experimental evidence shows that both FtsZ1 and FtsZ2 proteins are imported into chloroplasts and localized in the stroma (McAndrew et al. (2001) Plant Physiol. 127:1656-1666). The FtsZ1 and FtsZ2 proteins are reported to colocalize to rings at the plastid midpoint in Arabidopsis and other plants, where members of both families assemble into rings on stromal surface of the inner envelope membranes (Osteryoung, K W and McAndrew, R S (2001) Annu Rev Plant Physiol Plant Mol Biol 52:315-333; and McAndrew et al. (2001) Plant Physiol. 127:1656-1666). These FtsZ proteins have been characterized both biochemically and microscopically during non-photosynthetic bacterial division; efforts are under way to similarly characterize these proteins in plants. (for a review, see Osteryoung, K and McAndrew R S (2002) Annu Rev Plant Physiol Mol Biol 52: 315-322; and McAndrew et al. (2001) Plant Physiol. 127:1656-1666). A MinD protein has also been found encoded in plastid genomes of algae, as well as in the nuclear genomes of higher plants (Colletti K S, Tatersall E A, Pyke K A, Froelich A E, Stokes K D, Osteryoung K W (2000) Curr. Biol. 10:507-16,Moehs C P, Tian L, Osteryoung K W, DelaPenna D (2001) Plant Mol. Biol. In press); at least some of the MinD proteins include a cleavable chloroplast target sequence (Osteryoung, K and McAndrew R S (2002) Annu Rev Plant Physiol Mol Biol 52: 315-322). Reduced expression of MinD in Arabidopsis plants results in plants with asymmetrically constricted plastids (Colletti K S, Tatersall E A, Pyke K A, Froelich A E, Stokes K D, Osteryoung K W (2000) Curr. Biol. 10:507-16), suggesting that MinD also functions in plants to control the placement of the division ring to the center of the plastid. Both MinD as well as MinE are also encoded in the plastid genomes of unicellular algae (Wakasugi T, Nagai T, Kapoor M, Sugita M, Ito M, et al. (1997) Proc. Natl. Acad. Sci. USA 94:5967-72).
Currently, FtsZ, MinD, and MinE are the only obvious homologues of non-photosynthetic bacterial cell division genes known to exist in photosynthetic eukaryotes, and roles for MinE and MinD in plastid division have only recently been demonstrated, where they are involved in placement of the PD rings at the site of plastid constriction (Itoh et al. (2001) Plant Physiol. 127:1644-1655; Reddy et al. (2002) Planta. 215:167-176). Even the function of most of the other non-photosynthetic bacterial cell division proteins are not well understood, and they therefore cannot provide clues as to whether functional counterparts participate in plastid division. However, at least nine proteins localize to the division septum in E. coli (Margolin W (1198)Trends Microbiol. 6:233-38, Rothfield L I, Justice S S (1997) Cell 88:581-84), and the plastid division apparatus is likely to be at least as complex (Osteryoung K W, Pyke K A (1998) Curr Opin. Plant Biol. 1:475-79).
Therefore, there is a need to identify and characterize other genes involved in plastid division. The discovery of such genes is useful to further characterize the mechanism of plastid division. Moreover, these genes can then be manipulated to vary the number and size of plastids present in plant cells, in order to vary agronomic and horticultural characteristics of economically important plants, such as crop, ornamental, and woody plants.